TEMPORARY REMOVAL: Growing grass reduces soil N2O emissions and stimulates proliferation of denitrifying bacteria

Journal Pre-proof Growing grass reduces soil N2O emissions and stimulates proliferation of denitrifying bacteria

Haitao Wang, Lukas Beule, Huadong Zang, Birgit Pfeiffer, Shutan Ma, Petr Karlovsky, Klaus Dittert PII:

S0048-9697(20)30288-6

DOI:

https://doi.org/10.1016/j.scitotenv.2020.136778

Reference:

STOTEN 136778

To appear in:

Science of the Total Environment

Received date:

9 November 2019

Revised date:

9 January 2020

Accepted date:

16 January 2020

Please cite this article as: H. Wang, L. Beule, H. Zang, et al., Growing grass reduces soil N2O emissions and stimulates proliferation of denitrifying bacteria, Science of the Total Environment (2018), https://doi.org/10.1016/j.scitotenv.2020.136778

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© 2018 Published by Elsevier.

Journal Pre-proof Growing grass reduces soil N2O emissions and stimulates proliferation of denitrifying bacteria

Haitao Wanga, Lukas Beuleb, Huadong Zangc, Birgit Pfeifferd, Shutan Mae*, Petr Karlovskyb, Klaus Ditterta

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Department of Crop Science, Division of Plant Nutrition and Crop Physiology, University of Goettingen, Carl-Sprengel-Weg 1, 37075 Göttingen, Germany b

Molecular Phytopathology and Mycotoxin Research, Faculty of Agricultural Sciences, University of Goettingen, Grisebachstrasse 6, 37077 Göttingen, Germany College of Agronomy and Biotechnology, China Agricultural University, 100193, China

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c d

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Institute of Microbiology and Genetics, Department of Genomic and Applied Microbiology, Grisebachstrasse 8, 37077 Göttingen, Germany e

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[email protected]

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Corresponding author: Shutan Ma School of Environmental Science and Engineering Anhui Normal University Jiuhua south road 189 241002, Wuhu, China

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School of Environmental Science and Engineering, Anhui Normal University, Jiuhua south road 189, 241002, Wuhu, China

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The total number of words of the manuscript, including entire text from title page to figure legends: 7103 The number of words of the abstract: 283 The number of figures: 5 The number of tables: 1

Key words: soil N2O emissions, soil CO2 emissions, perennial ryegrass (Lolium perenne L.), soil bacteria, denitrification

Journal Pre-proof

Growing grass reduces soil N2O emissions and stimulates proliferation of denitrifying bacteria

Abstract

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Nitrogen (N) fertilization is the major contributor to nitrous oxide (N2O) emissions from agricultural soil. Previous studies have established that perennial grasses, barley, maize and other plants increase

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denitrification rates in soil. However, the majority of investigations of soil N2O emissions have been

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conducted without plants. The aim of this study, therefore, was to investigate how the presence of a

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perennial grass affects soil N2O emissions and denitrifier community size. To achieve this, a microcosm experiment was conducted with soil planted with perennial ryegrass (Lolium perenne L.) and bare soil,

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each with four levels of fertilizer (0, 5, 10 and 20 g N m−2; applied as calcium ammonium nitrate). The

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closed-chamber approach was used to measure soil N2O fluxes. Real-time PCR was used to estimate the biomass of bacteria and fungi and the abundance of genes involved in denitrification in soil. The results

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showed that the presence of ryegrass decreased the nitrate content in soil, as compared to bare soil,

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while no effect on dissolved organic carbon was observed. Cumulative N2O emissions of soil with grass were lower than in bare soil at 5 and 10 g N m−2 fertilization, but statistically indistinguishable in soil without fertilization and with 20 g N m−2. Fertilization did not affect the abundance of soil bacteria and fungi. Soil with grass showed greater abundances of bacteria and fungi, as well as microorganisms carrying narG, napA, nirK, nirS and nosZ clade I genes. It is concluded that growing perennial ryegrass can reduce soil N2O emissions and promote denitrifier community size compared to bare soil. This highlights the importance of plant cover for the reduction of N2O emissions from soil, particularly during N fertilization. Future research should explore how different plants species and communities affect denitrifying communities in soil.

Journal Pre-proof Key words: soil N2O emissions, soil CO2 emissions, perennial ryegrass (Lolium perenne L.), soil bacteria, denitrification

1. Introduction The increasing nitrous oxide (N2O) concentration in the atmosphere is among the most serious

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consequences of the anthropogenic alteration of the global nitrogen (N) cycle (Bakken and Frostegard, 2017). In addition to its high global warming potential and long atmospheric lifetime (IPCC, 2013), N2O

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has been shown to be the most important emitted compound involved in stratospheric ozone depletion

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(Ravishankara et al., 2009). The excessive input of mineral N into agricultural soils is one of the crucial

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factors contributing to soil N2O emissions. Denitrification is the predominant N2O producing biological

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process in soils (Bremner, 1997; Hu et al., 2015), which is strongly affected by the soil nitrate (NO3−) concentration (Köbke et al., 2018; Saggar et al., 2013). In the denitrification pathway, denitrifying

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microorganisms use NO3− as an electron acceptor and stepwise reduce it to gaseous N2. Incomplete denitrification results in the emission of gaseous intermediates like N2O. Therefore, conditions

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(Schlesinger, 2009).

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supporting complete denitrification is a possible approach to reducing N2O emissions from soil

Soil denitrification is regulated by enzymes such as NO3−, nitrite (NO2−) and N2O reductases that are produced by microorganisms. In arable soils, plant root architecture and exudation modifies soil structure, aeration and biological activity (Bertin et al., 2003; Kuzyakov and Xu, 2013), as well as soil microbial communities (Berg and Smalla, 2009). The majority of laboratory studies of soil N2O emissions, however, have not included plants, though it is known that growing plants increases denitrification activities in soil (Guyonnet et al., 2017; Klemedtsson et al., 1987). The inclusion of plants in laboratory experiments on N2O emissions may affect the outcome of these studies (Guyonnet et al., 2018a), as it has been estimated that 5% to 21% of all photosynthetically assimilated carbon (C) is released into the

Journal Pre-proof soil in the form of root exudates (Derrien et al., 2004; Nguyen, 2003). Consequently, the C turnover rate in the soil rhizosphere is estimated to be at least one order of magnitude greater than in the bulk soil (Kuzyakov, 2010). It has been suggested that root exudation will increase denitrification (Bijay-singh et al., 1988), as root-released C can serve as an electron donor (Philippot et al., 2007). Indeed, planted soils have a several times greater density of denitrifiers than unplanted soils (Chèneby et al., 2004; Herman et al., 2006). Growing perennial grasses, such as Festuca paniculata, Bromus erectus and Dactylis glomerata

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(Guyonnet et al., 2017), barley (Klemedtsson et al., 1987) and maize (Mahmood et al., 1997) has been

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shown to increase denitrification activities in soil. The stimulation of soil denitrification activity by plants depends on the plant species and soil water content (Bakken, 1988). Furthermore, root exudates have

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been shown to modulate soil microbial communities (Haichar et al., 2014, 2008).

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Ammonium (NH4+) uptake by plants has been shown to result in NH4+ depletion zones in the rhizosphere

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(Orcutt, 2000). In contrast, depletion zones of NO3− in the rhizosphere have been reported to be less pronounced, due to the high mobility of NO3− in most soils (Kuzyakov and Xu, 2013). The concentration

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of NO3− in soil, however, can rapidly decrease owing to uptake by plant roots (Tinker and Nye, 2000).

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Therefore, the availability of mineral N in soil is regarded as a major factor limiting denitrification

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(Philippot et al., 2007; Saggar et al., 2013). The response of soil N2O emissions to the application of mineral N fertilizer is exponential rather than linear (Shcherbak et al., 2014). Senbayram et al. (2012) reported that increasing the soil NO3− concentration resulted in a higher N2O/N2 ratio. The competition for NO3− between plants and denitrifiers can result in lower denitrification rates in planted soils (Qian et al., 1997). Similarly, regulation of denitrifying soil communities by NO3− has been reported from different ecosystems (Correa-Galeote et al., 2017; Deiglmayr et al., 2006; Enwall et al., 2005); however, the effect of the soil NO3− concentration on the abundance and diversity of denitrifiers remains to be determined. The main aim of this study was to investigate N2O emissions from soil with grass and bare soil under varying fertilizer levels. To achieve this, we used an incubation experiment with two experimental factors:

Journal Pre-proof soil planted with grass, and unplanted bare soil, each with four levels of N fertilizer addition. Soil N2O fluxes were determined using the closed-chamber approach. Real-time PCR (qPCR) assays were performed to estimate the biomass of soil bacteria and fungi, as well as microorganisms harboring genes involved in denitrification. We hypothesized that the presence of grass and the associated belowground modulations would (i) lower soil N2O emissions at each fertilizer level and (ii) promote the abundance of

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bacteria, fungi and denitrifiers, as compared to bare soil.

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2. Material and methods

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2.1 Soil collection

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Topsoil (0 to 25 cm) was collected from Reinshof agricultural research station (51°29'50.3"N,

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9°55'59.9"E), University of Goettingen, Lower Saxony, Germany [mean annual precipitation: 651 ± 24 mm; mean annual temperature: 9.2 ± 0.1°C (1981–2010, meteorological station at Goettingen, station ID:

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1691, Germany's Meteorological Service)]. The site had been cropped with winter oilseed rape (Brassica

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napus L.) (2015), winter wheat (Triticum aestivum L.) (2016), and winter barley (Hordeum vulgare L.) (2017) prior to soil collection on 23 March 2018. The soil was classified as Luvisol (IUSS, 2015) and the

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texture of the topsoil (0 to 25 cm) was composed of 61% silt, 23% sand and 16% clay. The bulk density was 1.3 g cm−3, the pH was 7.1  0.1, the soil total C concentration was 1.3% and the total N concentration was 0.13%. Following collection, the soil was stored in a polyvinyl chloride (PVC) container for three months at room temperature. Before incubation, the soil was air dried to 2% gravimetric water content and sieved through a 2-mm mesh. PVC cylinders (diameter: 20 cm; height: 20 cm) were used for incubation and sealed with removable lids (height: 5 cm) carrying butyl-rubber septa for headspace gas sampling. Soil moisture was adjusted to 35% water-filled pore space (WFPS) and soil was filled into the experimental pots in three layers of 3.7 cm each (11 cm in total) for manual compaction to a bulk density of 1.3 g cm−3, resulting in 4398 cm3 of air space for gas accumulation when the chambers were closed.

Journal Pre-proof The following day in a stepwise procedure, the soil was carefully irrigated and finally adjusted to 60% WFPS to avoid further soil compaction. During the incubation period, soil moisture was controlled to 60%  5% WFPS by weighing the pots and irrigating daily. 2.2 Experimental setup The experiment was conducted in a fully controlled climate chamber (Fitotron Walk in Plant Growth Room, Type SGR221 LED, Weiss Technik, Leics, UK). During the incubation period, the climate chamber

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was set to a light intensity of 520 µmol m–2 s–1 photosynthetically active photon flux density and a 25°C

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air temperature from 06:00 to 22:00 as ‘day mode’ (16 h), and from 22:00 to 06:00 (8 h) as ‘night mode’

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with no light and a 12°C air temperature. The large temperature discrepancy was set in order to reduce

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nighttime respiration and preserve photosynthetic C assimilates.

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The experiment consisted of two groups: soil with perennial ryegrass (Lolium perenne L.) (DSV AG, Salzkotten, Germany), and bare soil. Each had four different fertilizer levels (0, 5, 10 and 20 g N m−2),

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resulting in a total of eight treatments. Each treatment was performed in triplicate, yielding a total of 24 pots. Before the first sampling date, grass was sown at a density of approximately 5000 seeds m −2 and

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pre-incubated for four weeks to allow grass establishment in the pots. The treatments with bare soil

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were treated equally but without plant cultivation. Calcium ammonium nitrate N fertilizer [76% ammonium nitrate (NH4NO3) and 24% calcium carbonate (CaCO3)] was applied after dissolution in distilled water. Half of the total N fertilizer was applied after the first collection of soil and gas samples on day 1 (3 August 2018); the other half was applied after 28 days, with a full measuring period of 56 days. Two days before the first fertilization, the grass was cut to a height of 4 cm. Following this, the grass was cut every two weeks and the shoot dry matter was determined from air-dried material. At the end of the experiment, the roots were collected as well. Roots were carefully washed, air-dried and weighed. The

Journal Pre-proof total C and N of finely ground dry grass shoots and roots were determined on an NA-1500 N elemental analyzer (Carlo Erba, Milano, Italy). Grass N uptake and C assimilation were calculated as (1) and ,

(2)

(

)

(

)

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( )

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where DM refers to the dry matter of the harvested grass. Apparent N recovery (ARN) was calculated as .

(3)

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Gas samples of 25 mL in volume were collected using a syringe inserted in the headspace of sealed lids.

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Samples were directly transferred to a pre-evacuated 12-mL Exetainer vial (Labco, Lampeter, UK). Gas

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samples were collected at 0, 20 and 40 min after the pots were sealed. In the first week after each fertilization, gas samples were collected every day to capture the fertilization-induced peaks. In the

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following three weeks, gas samples were taken at larger intervals of 2 to 4 days.

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Soil samples were taken on day 0 (one day before the first fertilization), day 7, day 14, day 28 (before the

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second fertilization), day 35 and day 56 (final collection of samples). In order to avoid the disturbance of soil structure by soil sampling during the incubation period, the soil samples were collected from two other parallel spare pots per gas sampling pot (on day 0, day 7 and day 14, soil samples were only collected from the first spare pot; on day 35 and day 56, they were only collected from the second spare pot), which were placed alongside the gas sampling pots. Only the last soil samples were taken from the pots on which gas measurement was performed. The soil was taken using a 16-mm-diameter auger. Remaining holes were filled with reagent glasses. Approximately 60 g of fresh soil was taken and sieved through a 2-mm mesh. The samples were homogenized and divided into six subsamples for soil NH4+ and NO3−, soil pH, total C and N, and dissolved organic carbon (DOC) analyses, as well as DNA extraction and WFPS determination.

Journal Pre-proof 2.3 Gas and soil sample analysis Gas samples were analyzed on an Agilent 7890A gas chromatograph (Agilent Technologies, Santa Clara, USA) equipped with a thermal conductivity detector for the determination of carbon dioxide (CO2) concentrations and an electron capture detector for the determination of N2O concentrations. The flux rates (CO2 and N2O) were calculated using a linear or nonlinear regression of the gas concentration over time (Parkin et al., 2012; Wang et al., 2013). Cumulative emissions were calculated by interpolation.

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To determine soil NH4+ and NO3− concentrations, subsamples (10 g) of sieved fresh soil were extracted by

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adding 50 mL of 0.0125-M CaCl2. Mixtures were shaken for one hour, filtered (MN615 ¼; pore size: 4–12

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µm; Macherey-Nagel, Düren, Germany) and subsequently stored at −20°C until analysis. NH4+ and NO3−

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concentrations in the extracts were determined using a San++ continuous flow analyzer (Skalar Analytical, Breda, The Netherlands). Soil pH was measured from 10 g of air-dried soil dissolved in 50 mL of 0.01-M

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CaCl2 solution using a pH meter (Sartorius, Göttingen, Germany). Total C and N measurements were

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performed with finely ground air-dried soil using an NA-1500 elemental analyzer (Carlo Erba, Milano, Italy). Prior to the measurement of total C and N, the air-dried soil was fumigated in a hydrogen chloride

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(HCl) atmosphere using 3-M HCl for one week to remove carbonates (Harris et al., 2001). For DOC

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measurements, 10 g of fresh soil was extracted using 40 mL of 0.5-M K2SO4 (potassium sulfide). The solution was shaken for two hours and filtered (MN615 ¼; pore size: 4–12 µm; Macherey-Nagel, Düren, Germany). Extracts were stored at −20°C until determination of total C and inorganic C concentrations using a TOC/TIC analyzer (Multi C/N 2100, Analytik Jena, Jena, Germany). 2.4 DNA extraction from soil and qPCR Total DNA extraction from 50 mg of freeze-dried and finely ground soil material was performed in accordance with Beule et al. (2019). The quality and quantity of DNA were examined on 0.8% (w/v) agarose gels stained with ethidium bromide. The samples were diluted at a ratio of 1:50 in doubledistilled water and stored at −20°C. qPCR was conducted in a CFX384 Thermocycler (Biorad, Rüdigheim,

Journal Pre-proof Germany). qPCR reactions for bacterial 16S rRNA and fungal 18S rRNA genes, as well as the denitrification genes narG and napA for NO3− reduction, nirK and nirS for NO2− reduction, and nosZ clade I and II for N2O reduction, were performed as described previously by Beule et al. (2019). 2.5 Statistical analysis All data were tested for homogeneity of variances (Levene’s Test) and normal distribution (Shapiro–Wilk test). Differences among treatments of cumulative data (N uptake and C assimilation by grass and

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cumulative CO2 and N2O emissions) or data without repeated measurements (soil bacteria, fungi and

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denitrifiers) were assessed by performing a t-test or one-way ANOVA with Tukey's honestly significant

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difference (HSD) post-hoc test for parametric data, or the Mann–Whitney-U test or Kruskal–Wallis test

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with multiple comparison extension for non-parametric data. Differences among treatments of repeatedly measured data (DOC, NO3−, NH4+, CO2 and N2O fluxes) were analyzed using linear mixed effect

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(LME) models. In the models, either the fertilizer level or the treatment of bare soil versus soil with grass

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were set as a fixed effect, and sampling date and replicate pot set as random effects. The data were partially log10- or square-root-transformed to meet the criteria for an LME model. Statistical significance

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was considered as p < 0.05, with marginal statistical significance at p < 0.1. All statistical analyses were

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performed in R version 3.5.2 (R Core Team, 2018).

3. Results 3.1 Grass N uptake and C assimilation In soil with grass, total plant N uptake, which was calculated by the dry matter of grass shoots and roots, ranged from 6.5 to 13.9 g N m−2 in fertilized pots, compared to 4.1 g N m−2 in the unfertilized treatment. The ARNs of fertilized treatments were 50% ± 2%. Plant shoot N uptake at the 10 g N m−2 fertilizer level was lower than that at 20 g N m−2 (p = 0.006) and greater than those at 0 and 5 g N m−2 (p < 0.01) (Tab. 1).

Journal Pre-proof The total N uptake throughout the incubation period increased along with increasing fertilizer application (p < 0.03) (Tab. 1). Shoot C, root C and total C assimilation were greater in fertilized than unfertilized pots (p < 0.04) (Tab. 1), but did not differ among the 5, 10 and 20 g N m −2 fertilization treatments. 3.2 Soil DOC, NO3− and NH4+ dynamics during incubation DOC was slightly increased in the first two weeks, and gradually decreased in the following weeks in both

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bare soil and soil with grass (Fig. 1A, B). DOC concentrations did not differ among bare soil and soil with

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grass, nor among fertilizer levels (Fig. 1A, B). Soil NO3−-N content in bare soil was always greater than in

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soil with grass at all fertilizer levels (p < 0.001) (Fig. 1C, D). Fertilization (calcium ammonium nitrate) led

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to increased NO3−-N concentrations in bare soil (p < 0.05) (Fig. 1C). Compared to the background NO3− (approximately 2 g of NO3−-N m−2) in unfertilized bare soil, soil NO3−-N was close to zero in the

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unfertilized treatment of soil with grass (Fig. 1D). Furthermore, in soil with grass, at the 20 g N m−2

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fertilizer level, soil NO3−-N was greater than at all other fertilizer levels (p < 0.05) (Fig. 1D). When N

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fertilizer was applied, NH4+-N was marginally greater in bare soil than in soil with grass (p < 0.1) (Fig. 1E,

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3.3 CO2 and N2O emissions

The presence of grass strongly enhanced CO2 emissions compared to bare soil, especially in treatments with fertilizer (p < 0.003) (Fig. 2A). Fertilization had no effect on CO2 emissions (neither on short-term rates nor on cumulative fluxes) in bare soil (Fig. 2A). In soil with grass, however, CO2 emission rates and cumulative fluxes were increased by fertilizer application (p < 0.001) (Fig. 2B). In contrast to CO2 fluxes, N2O emissions from bare soil were greater than from soil with grass at each fertilizer level (p < 0.05) (Fig. 3). At the 5 and 10 g N m−2 fertilizer levels, bare soil showed greater cumulative N2O emissions than soil with grass (p < 0.05). Cumulative N2O emissions from soil with grass at the 0, 5 and 10 g N m−2 fertilizer levels were lower than at 20 g N m−2 (p < 0.05) (Fig. 3B).

Journal Pre-proof 3.4 Soil microbial gene abundances in bare soil and soil with grass At the end of the experiment (day 56), the abundances of bacteria, fungi and denitrification genes in soil were quantified. The fertilization rate did not affect the abundances of bacteria, fungi and denitrification genes (Fig. 4, Fig. 5). At the 0 and 20 g N m−2 fertilizer levels, the soil with grass showed marginally greater bacterial 16S rRNA gene copy numbers than bare soil (p < 0.08) (Fig. 4A). Similarly, the number of fungal 18S rRNA gene copies did not differ among fertilizer levels, but were greater in soil with grass than

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bare soil at the 0, 10 and 20 g N m−2 fertilizer levels (p < 0.05) (Fig. 4B). The abundance of narG was

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greater in the soil with grass than bare soil at the fertilizer level of 20 g N m−2 (p < 0.005) (Fig. 5A). At the

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0 and 5 g N m−2 fertilizer levels, gene copy numbers of napA in soil with grass were greater than in bare soil (p < 0.06) (Fig. 5B). At each fertilizer level, nirK gene copy numbers were greater in soil with grass

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than in bare soil (p < 0.09) (Fig. 5C). The abundance of nirS was increased in soil with grass compared to

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bare soil when 5, 10 or 20 g N m−2 of fertilizer was applied (p < 0.1) (Fig. 5D). At the fertilization rate of 20

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g N m−2, nosZ clade I gene copies were marginally greater in soil with grass than in bare soil (p < 0.07) (Fig. 5E). No differences between soil with grass and bare soil at any fertilizer level were detected for nosZ

4. Discussion

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clade II (Fig. 5F).

4.1 Soil organic C turnover and CO2 emissions The soil microbial community is the main driver of soil respiration and organic C mineralization in bare soils (Liu et al., 2018; Li et al., 2018). The slight increase in DOC in the first two weeks may have been due to the rewetting of the dry soil to 60% WFPS (Kalbitz et al., 2000). For example, when Lundquist et al. (1999) exposed soil to wet–dry cycles, soil aggregates were partly decomposed and their C was found in the DOC fraction. In the first three weeks of the experiment, soil CO2 emissions increased gradually in bare soils, indicating a recovery of the microbial respiration from the rewetted air-dried soil (Fig. 2A). As

Journal Pre-proof the soil was already pre-incubated for four weeks before the application of fertilizer, this may be seen as indication that the recovery of the soil microbial activity in the bare soil may take approximately seven weeks under the given conditions. One reason for this long recovery period in bare soil may be the limitation of available C. Three weeks after the first fertilization, the stable CO2 emissions and slow DOC consumption rate may point towards a stabilized soil microbial community after three weeks. Pausch and Kuzyakov (2018) reviewed the distribution of C compounds in soil that were released by

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roots. They concluded that 12% of the assimilated C is emitted from the plant as root-derived CO2 and 5%

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is deposited in the rhizosphere. Most plant root exudates have been reported to be readily available to

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soil microorganisms since they can be metabolized within a few hours (Jones et al., 2005; Kuzyakov and Xu, 2013). Moreover, it is reasonable to expect a greater DOC content in soils with grass then in bare soil

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given the estimation that 5% of the assimilated C is sequestrated in the rhizosphere (Pausch and

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Kuzyakov, 2018). However, no such increase was found in our study. Zhang et al. (2018) reported that

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heavy grazing lowered the C input and decreased C accumulation and total soil organic C contents, due to reduced aboveground tissue (Schönbach et al., 2011), more exposure of the soil surface, and thus

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increased loss of soil moisture (Zhao et al., 2007) and stimulated compensatory growth of new leaves

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(Zhao et al., 2008). Therefore, the intensive cutting throughout our experiment likely contributed to the lack of increased soil DOC.

4.2 Soil mineral N and N2O emissions The time series of N2O emissions followed a pattern that was similar to that for soil NO3− concentrations. For example, in each of the eight treatments, the second N2O emissions peak, which followed the second fertilizer application, was greater than the first peak (Fig. 3A and 3B). Additionally, in contrast to soil with grass, bare soil treatments showed considerably greater N2O emissions, which lasted over the entire study period. This was further associated with decreasing soil NO3− concentrations, which, in contrast to treatments in bare soil, did not reach zero after 56 days. In soil with grass, N2O emissions fell to nearly

Journal Pre-proof zero two weeks after each fertilizer application, which was associated with exploited soil NO3−. The relationships between soil NO3− and N2O emissions indicate that, under these conditions, soil NO3− is the predominant factor controlling soil N2O emissions (Dong et al., 2018; Ji et al., 2018; Zhou et al., 2017). At 60% WFPS, both nitrification and denitrification are expected to be important contributors to soil N2O emissions, as this moisture level is seen as the threshold between aerobic and anaerobic conditions (Menéndez et al., 2012; Volpi et al., 2017). We anticipate that, in the first week after fertilization, both

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pathways (nitrification and denitrification) contributed to the observed N2O emissions. In the following

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weeks, denitrification likely became the predominant process contributing to N2O emissions owing to

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NH4+ removal (Fig. 1E, 1F) and lowered oxygen partial pressure induced by root O2 consumption

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(Klemedtsson et al., 1987).

Due to root activity, two opposing effects on denitrification will occur: (i) oxygen consumption by aerobic

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root activity (root respiration consuming O2); and (ii) plant transpiration, leading to drainage of coarse

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soil pores and thus increased air-filled pore space, and hence increased oxygen availability. In our study, although water content was adjusted every 1–2 days, soil moisture likely fell to levels below 55% WFPS

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before irrigation due to plant transpiration. Therefore, lower soil moisture due to plant transpiration

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may increase oxygen diffusion into the soil and thereby suppresses denitrification (Menéndez et al., 2012; Volpi et al., 2017). Many previous studies have reported that the presence of plants increases denitrification (Guyonnet et al., 2017; Klemedtsson et al., 1987; Mahmood et al., 1997). However, our study was not a perfect proof of the opposite, but at least provides evidence that the earlier reported promotion of denitrification does not always happen; at least, plants do not always induce higher N2O emissions. 4.3 Influence of the presence of plants on soil microbial abundances The population size and diversity of microbial communities have repeatedly been shown to increase in the presence of plants (Guyonnet et al., 2018b; Haichar et al., 2008; Li et al., 2018). In line with this, our

Journal Pre-proof results showed that population densities of both soil bacteria and fungi increased with the presence of ryegrass. Considering C-limited conditions in unplanted soil, we assume that root-derived input of easily available C promoted these microbial populations. Plant root exudates are known to modulate both microbial biomass and community composition (Benizri et al., 2007; Henry et al., 2008; Langarica-Fuentes et al., 2018). However, a limited number of studies have explored how plants influence genes involved in denitrification (Henry et al., 2008; Pivato et al.,

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2017). We found that, with the exception of nosZ clade II, all denitrification genes were promoted in the

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presence of ryegrass, which may be due to the root exudation of easy available C. Graf (2015) proposed

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a greater affinity of nosZ clade I-carrying microorganisms to root exudates than for those carrying nosZ clade II. Our findings agree with the suggestions of Graf (2015): there was a trend that nosZ clade I genes

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were greater in soil with grass, whereas nosZ clade II showed no preference for bare versus planted soil.

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4.4 Relationship of reduced N2O emissions and increased denitrifying gene abundances in soil with

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grass

In our study, N2O emissions were reduced even though denitrification genes increased under grass. We

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found that the emission factors of bare soil and soil with grass were 1.4%–1.8% and 0.5%–0.8%,

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respectively, which corresponded to a 50% plant N uptake, indicating that the role of plant N uptake in reducing the substrate for N2O emissions has been underplayed. It should be mentioned that, due to the limitation of the experimental design, soil samples and gas samples were not taken from the same pots, and N2O emissions were highly variable. Therefore, it was not possible to correlate N2O emissions and NO3− concentrations in this study. In soil with grass at the 20 g N m−2 fertilizer level, soil NO3− concentrations were not depleted by plant uptake and, concurrently, cumulative N2O emissions at this level were more than twice as high as those at the 10 g N m−2 fertilizer level, suggesting that an N input that exceeds the plant’s needs can exponentially increase soil N2O emissions. Previous field studies have shown congruent results

Journal Pre-proof (Groenigen et al., 2010; Philibert et al., 2012; Shcherbak et al., 2014). This confirms findings that soil with ryegrass has a greater potential for N2O emissions when NO3− is not limiting denitrifier activity (Hayashi et al., 2015; Philippot et al., 2013; Uchida et al., 2011). The much lower cumulative N2O emission levels in soil with grass, as compared to bare soil, at the 5 and 10 g N m−2 fertilizer levels, was most likely due to plant uptake of soil mineral N. It was recently suggested that soil NO3− availability affects denitrifying communities (Deiglmayr et al.,

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2006; Saggar et al., 2013; Tang et al., 2016). However, we found no link between fertilizer level and

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denitrification-related genes. The reason might be the rapid and almost complete utilization N from the

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soil by ryegrass, resulting in low soil NO3− concentrations at all fertilization levels. It should be noted, however, that this observation requires further study, since denitrifiers were only quantified at the end

re

of our experiment. Therefore, potential changes in microbial communities during the course of our

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experiment may have remained undetected. Further studies should include C substrate amendment as

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5. Conclusion

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well as repeated determination of denitrifier population size using a non-destructive soil sampling design.

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In our incubation experiment of soil planted with ryegrass and bare soil under different N fertilizer levels, we found that, although the proliferation of denitrifying bacteria in soil with grass was stimulated, N2O emissions were still lower, together with large amounts of plant N uptake. The results infer that soil mineral N is more related to N2O emissions than soil denitrifying genes. However, because of the higher potential of denitrification in soil with grass, the risk of high N2O emissions should also be noted, especially when N fertilizer exceeds the requirements of plants. Future studies should focus on how different plant species and their root exudates affect soil N2O emissions and related soil microorganisms. It is necessary to integrate plants, soil N fertilizer and soil microorganisms together to understand the mechanisms of N2O production and its mitigation.

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6. Acknowledgements This project was supported by the China Scholarship Council. We also thank Susanne Koch, Simone Urstadt, Marlies Niebuhr, Ulrike Kierbaum, and Jürgen Kobbe for diligent and skillful assistance.

References

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Bakken, L.R., Frostegard, A., 2017. Sources and sinks for N2O, can microbiologist help to mitigate N2O emissions? Environ. Microbiol. 19, 4801–4805. https://doi.org/10.1111/1462-2920.13978

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Zhou, M., Zhu, B., Wang, X., Wang, Y., 2017. Long-term field measurements of annual methane and nitrous oxide emissions from a Chinese subtropical wheat-rice rotation system. Soil Biol. Biochem. 115, 21–34. https://doi.org/10.1016/j.soilbio.2017.08.005

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Table 1. Total N uptake and C assimilation of grass shoots and roots throughout the experimental period (56 days) at each fertilizer level in soil with grass. -2

Fertilizer level (g

-2

N uptake (g N m )

C assimilation (g C m )

-2

Nm )

Root

Total

Shoot

Root

Total

0

2.6 ± 0.1c

1.5 ± 0.2b

4.1 ± 0.1d

58.3 ± 2.6b

58.7 ± 2.8b

117.0 ± 4.4b

5

4.7 ± 0.2c

1.8 ± 0.1a

6.5 ± 0.2c

98.4 ± 4.3a

10

7.4 ± 0.7b

2.0 ± 0.1a

9.3 ± 0.5b

20

11.8 ± 0.7a

2.1 ± 0.1a

13.9 ± 0.5a

ro

of

Shoot

re

-p

115.8 ± 5.3a 123.6 ± 9.2a

74.3 ± 0.9a

172.6 ± 4.7a

74.8 ± 4.1a

190.5 ± 1.2a

68.1 ± 2.6a

191.8 ± 10.8a

Means ± standard errors followed by different lowercase letters indicate significant differences among

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fertilizer levels within each parameter (one-way ANOVA with Tukey’s HSD test or Kruskal-Wallis test with

Jo

ur

na

multiple comparison extension) at p < 0.05.

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Bare Soil 8

Fertilization level (g N m-2) 20 10 5 0

A

7

DOC (g C m-2)

Soil with grass

6 5

B

4 3

D

ro

C

12

-p

10 8 6

re

NO3- (g N m-2)

14

4

lP

2 0

F

na

E

0.8

ur

0.6 0.4

Jo

NH4+ (g N m-2)

of

2

0.2 0.0 0

10

20

30

40

Sampling dates

50

0

10

20

30

40

50

Sampling dates

Fig. 1. Time course of dissolved organic carbon (DOC) concentrations in (a) bare soil and (b) soil with grass, and soil NO3--N content in (c) bare soil and (d) soil with grass, and soil NH4+-N concentrations in (e) bare soil and (f) soil with grass during the 56 days growing period. Solid lines with points of different gray intensities represent different fertilizer levels (0, 5, 10, and 20 g N m-2), dashed vertical lines indicate fertilization dates (day 1 and 28). Error bars represent the standard error of the mean (n = 3).

Journal Pre-proof Cumulative CO2-C emission

CO2-C fluxes Fertilization level (g N m-2)

4.5 3.5

300 a

a

a

a

2.5

200 100

-2

-1

CO2 (g C m d )

400

20 10 5 0

-2

A Bare soil

* * * *

1.5

of

B Soil with grass (incl. root + shoot respiration)

a

a

a

0 400 300

ro

12 10 8 6 4 2 0

CO2 (g C m )

5.5

b

10

20

30

40

lP

0

re

-p

200

0 50

0

5

10

20

Fertilization level (g N m-2)

na

Sampling dates

100

Fig. 2. CO2 emission dynamics and cumulative CO2 emission during the growing period (56 days) from

ur

bare soil (A) and soil with grass (B). Error bars represent the standard error of the mean of each

Jo

treatment (n = 3). Solid lines with points of different gray intensities represent different fertilizer levels (0, 5, 10 and 20 g N m-2). Dashed vertical lines indicate fertilization dates (day 1 and 28). Asterisks indicate significant differences in cumulative CO2 emission between bare soil and soil with grass at the same fertilizer level (T-test or Mann-Whitney-U test), lowercase letters indicate significant differences in cumulative CO2 emission among fertilizer levels within bare soil or within soil with grass (one-way ANOVA with Tukey’s HSD test or Kruskal-Wallis test with multiple comparison extension) at p < 0.05.

Journal Pre-proof Cumulative N2O-N emission a

A Bare soil

-2

Fertilization level (g N m ) 20 10 5 0

20

200

10

a

0

B Soil with grass

a

100

* *

0

of

40

300

a

30

400

ro

-2

-1

N2O (mg N m d )

30

-p

20

0 10

20

30

40

lP

0

re

10

a

300 200

b

b

0

5

100

b

0 50

10

20

Fertilization level (g N m-2)

na

Sampling dates

400

-2

40

N2O (mg N m )

N2O-N fluxes

ur

Fig. 3. N2O emission dynamics and cumulative N2O emission during the growing period (56 days) from bare soil (A) and soil with grass (B). Error bars represent the standard error of the mean of each

Jo

treatment (n = 3). Solid lines with points of different gray intensities represent different fertilizer levels (0, 5, 10 and 20 g N m-2). Dashed vertical lines indicate fertilization dates (day 1 and 28). Asterisks indicate significant differences in cumulative N2O emission between bare soil and soil with grass at the same fertilizer level (T-test or Mann-Whitney-U test), lowercase letters indicate significant difference in cumulative N2O emission among fertilizer levels within bare soil or within soil with grass (one-way ANOVA with Tukey’s HSD test or Kruskal-Wallis test with multiple comparison extension) at p < 0.05.

3e+9

A 2e+9

Bare Soil Soil with grass

*

†

1e+9

B

* *

-p

3e+8

*

re

2e+8

ro

of

0 4e+8

1e+8

0 5

10

20

na

0

lP

Fungal 18S rRNA gene copy number g-1 dry soil

Bacterial 16S rRNA gene copy number g-1 dry soil

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Fertilization level (g N m-2)

ur

Fig. 4. Bacterial 16s rRNA (A) and fungal 18s rRNA (B) gene copy number per g dry soil in bare soil and

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soil with grass under different fertilizer levels (0, 5, 10 and 20 g N m-2) at the end of the growing period (days 56). Error bars represent standard error of the mean of each treatment (n = 3). Asterisks denote differences between bare soil and soil with grass (* p < 0.05), Daggers represent marginal differences between bare soil and soil with grass († p < 0.1).

B

Bare Soil Soil with grass

1e+9

*

5e+8

1e+9

†

*

5e+8

†

*

†

*

†

*

†

lP

E

F

2e+7

†

2e+7

na

2e+7

1e+7

5e+6

5

Jo

0

10

2e+8

1e+8

re

1e+8

2e+7

3e+8

-p

2e+8

of

D

1e+7

5e+6 20

0

5

10

nirS gene copy number g-1 dry soil

C

ro

3e+8

0

ur

nosZ clade I gene copy number g-1 dry soil

nirK gene copy number g-1 dry soil

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napA gene copy number g-1 dry soil

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nosZ clade II gene copy number g-1 dry soil

narG gene copy number g-1 dry soil

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Fertilization level (g N m-2)

Fig. 5 narG (A), napA (B), nirK (C), nirS (D), nosZ clade I (E) and nosZ clade II (F) gene copy number per g dry soil in bare soil and soil with grass under different fertilizer levels (0, 5, 10 and 20 g N m-2) at the end of the growing period (days 56). Error bars represent standard error of the mean of each treatment (n = 3). Asterisks denote differences between bare soil and soil with grass (* p < 0.05). Daggers represent marginal differences between bare soil and soil with grass († p < 0.1).

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Declaration of interests ☒ The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. ☒The authors declare the following financial interests/personal relationships which may be considered as potential competing interests:

Graphical abstract

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About 50% of fertilized N was recovered in growing grass after two months of incubation N2O emissions were lower in soil with grass than bare soil at medium fertilizer levels (5 and 10 g N m-2) Growing grass stimulated the proliferation of almost all the denitrifying bacteria except nosZ clade II

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